FungiJournal of

Article

Targeted Delivery of Gene Silencing in Fungi Using GeneticallyEngineered Bacteria

Jonatan Niño-Sánchez 1,2 , Li-Hung Chen 1,3 , Jorge Teodoro De Souza 1,4, Sandra Mosquera 1,5

and Ioannis Stergiopoulos 1,*

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Citation: Niño-Sánchez, J.;

Chen, L.-H.; De Souza, J.T.;

Mosquera, S.; Stergiopoulos, I.

Targeted Delivery of Gene Silencing

in Fungi Using Genetically

Engineered Bacteria. J. Fungi 2021, 7,

125. https://doi.org/10.3390/

jof7020125

Received: 29 December 2020

Accepted: 5 February 2021

Published: 9 February 2021

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1 Department of Plant Pathology, University of California Davis, Davis, CA 95616, USA;jnino@ucdavis.edu (J.N.-S.); lhchen@ucdavis.edu (L.-H.C.); jorge.souza@dfp.ufla.br (J.T.D.S.);smosquera@ucdavis.edu (S.M.)

2 Department of Microbiology and Plant Pathology, University of California Riverside, Riverside,CA 92521, USA

3 Department of Plant Pathology, National Chung-Hsing University, Taichung 40227, Taiwan4 Department of Plant Pathology, Federal University of Lavras (UFLA), Lavras, MG 37200-000, Brazil5 Department of Ciencias Biológicas, Universidad EAFIT, Medellín 050022, Colombia* Correspondence: istergiopoulos@ucdavis.edu; Tel.: +1-530-400-9802

Abstract: Exploiting RNA interference (RNAi) in disease control through non-transformative meth-ods that overcome the hurdle of producing transgenic plants has attracted much attention over the lastyears. Here, we explored such a method and used non-pathogenic bacteria as a versatile system fordelivering RNAi to fungi. Specifically, the RNaseIII-null mutant strain of Escherichia coli HT115(DE3)was transformed with two plasmid vectors that enabled the constitutive or IPTG-inducible pro-duction of double-stranded RNAs (dsRNAs) against genes involved in aflatoxins production inAspergillus flavus (AflC) or virulence of Botrytis cinerea (BcSAS1). To facilitate the release of the dsR-NAs, the bacterial cells were further genetically engineered to undergo a bacteriophage endolysinR-mediated autolysis, following a freeze-thaw cycle. Exposure under in vitro conditions of A. flavusor B. cinerea to living bacteria or their whole-cell autolysates induced silencing of AflC and BcSAS1 ina bacteria concentration-dependent manner, and instigated a reduction in aflatoxins production andmycelial growth, respectively. In planta applications of the living bacteria or their crude whole-cellautolysates produced similar results, thus creating a basis for translational research. These resultsdemonstrate that bacteria can produce biologically active dsRNA against target genes in fungi andthat bacteria-mediated RNAi can be used to control fungal pathogens.

Keywords: RNA interference; HIGS; SIGS; dsRNA; cross-kingdom RNAi; Escherichia coli HT115(DE3);Botrytis cinerea; Aspergillus flavus; aflatoxins; bacterial autolysis

1. Introduction

RNA interference (RNAi) is a conserved mechanism of gene-silencing in nature withimmense therapeutic potential. It is triggered by inverse transcripts that arecomplementaryto the mRNA of a target gene, including double-stranded RNAs (dsRNAs) and small-interfering RNAs (siRNAs) that are produced by the DICER-mediated cleavage of dsRNAsinto small pieces of 19–25 nucleotides [1]. Since its discovery, RNAi has been transformativeto the study of gene function in numerous eukaryotic organisms and has created immenseopportunities for its exploitation in medicine and disease control [2,3]. RNAi therapeuticsis currently a rapidly evolving biotechnological field and despite the many challengesencountered along the way, 2018 marked a new era with the approval by the US Food andDrug Administration (FDA) of the first RNAi-based drug [2,4]. At present, several RNAi-based drugs that target diverse and rare human diseases are currently under developmentor undergoing phase I or phase II clinical trials. In addition, much effort is placed on theconcomitant development of delivery strategies to target cells and tissues, which remains akey issue in the exploitation of RNAi therapeutics [2,5,6].

J. Fungi 2021, 7, 125. https://doi.org/10.3390/jof7020125 https://www.mdpi.com/journal/jof

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In crop protection, RNAi has mostly been utilized in the form of host-induced genesilencing (HIGS), an approach in which plants are genetically engineered to producedsRNAs or siRNAs against genes in target pests and pathogens [7]. Several studies havehighlighted the utility of HIGS in a variety of crops for the control of a wide-range of insectpests, nematodes, viruses, and fungal plant pathogens [7–10]. HIGS has also proven to bean effective method for reducing mycotoxin contamination in food and feed, including ofthe notorious aflatoxins [8,11–15]. Aflatoxins are secondary metabolites produced mainlyby species of Aspergillus spp. and particularly by A. flavus and A. parasiticus [16,17]. At least20 different aflatoxins have been described but aflatoxin B1 (AFB1), B2 (AFB2), G1 (AFG1)and G2 (AFG2) are the four most common types, with AFB1 and AFB2 mainly produced byA. flavus, and AFG1 and AFG2 by A. parasiticus [16,17]. They are highly toxic mycotoxins,well-known for their carcinogenicity and other serious illnesses related to liver damage andimmunosuppression that they can cause in humans and livestock [18]. They enter the foodchain mainly through contaminated agricultural commodities, with grain and nut cropssuch as maize, rice, and peanuts being major sources of contamination [17]. Estimates bythe Food and Agriculture Organization (FAO) are that over 25% of the global food supplyis contaminated by mycotoxins, including aflatoxins, whereas according to the WorldHealth Organization (WHO), approximately 4.5 billion people are chronically exposedthrough their diet to these poisonous compounds, with often detrimental effects [16,17].Thus, finding effective ways to mitigate contamination of food and feed by aflatoxinsis crucial.

Despite the potential of HIGS for disease control, off-target effects, difficulties in planttransformation, and low public acceptance of transgene technology have restricted itsapplication in the field [7,15,19,20]. Virus-induced gene silencing (VIGS), which exploitsrecombinant viruses as vectors to deliver RNAi molecules to plants, is an alternativeto HIGS but to date it has mostly been utilized as a functional tool to study genes ofinterest in different crops, rather than for disease control [21–23]. The relatively recentdiscovery that externally applied dsRNAs and siRNAs can be taken-up by fungal cellsfrom their environment and induce silencing in the transfected cells, has attracted muchattention for its potential use as an alternative to HIGS for delivering RNAi to targetpathogens [24]. Thus, spray-induced gene silencing (SIGS) has recently emerged as anovel, environmentally-friendly disease control method. At present, several attempts togenerate RNAi-based biopesticides that utilize synthetic dsRNAs or siRNAs against genesin target pests and pathogens are underway [25,26]. Such RNA-based biopesticides canbe customized to be species-specific or to target a broad range of pathogens, even acrossdifferent taxonomic groups. Moreover, multiple genes can be simultaneously silenced, thusreducing the risk for resistance development [25–27]. However, a major limitation to theuse of synthetic RNAi molecules as biopesticides is the cost of their large-scale productionand formulation as well as the limited stability of the naked RNAi molecules under fieldconditions [26,27]. To overcome these limitations, a variety of synthetic materials have beendeveloped to encapsulate and protect dsRNA from nuclease degradation. These include adouble-layered hydroxide clay nanosheet that has been proved effective to protect dsRNAtargeted against some viral diseases, or carbon nanotubes that successfully deliver siRNAand trigger posttranscriptional gene silencing in plant cells. Nevertheless, the use of thesesynthetic materials is still limited [3,28].

Here, we explored the possibility of using non-pathogenic bacteria as a versatileproduction and delivery system of RNAi to fungi. Bacteria-mediated delivery of RNAihas been successfully used to silence genes in Caenorhabditis elegans [29], insects [27],shrimps [30], and mammalian cells [31] but the utility of this approach for the control offungi, to our knowledge, has not been exploited yet [26]. In our approach, a mutant ofEscherichia coli lacking RNaseIII (an enzyme that degrades dsRNAs in bacterial cells) [29]was genetically engineered to express dsRNAs against A. flavus [32,33], and Botrytis cinerea,a notorious plant pathogen with a remarkably broad host range [34], and to further undergoa programmable autolysis [35], thus allowing the destruction of the bacterial cells and the

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release of the fungicidal dsRNAs. Our data show that the engineered bacteria accumulatedhigh levels of dsRNAs, whereas subsequent treatment under in vitro or in planta conditionsof A. flavus and B. cinerea with the living bacteria or the induced bacterial autolysatesresulted in silencing of the complementary fungal genes and a reduction in production ofaflatoxins or fungal growth, respectively.

2. Materials and Methods2.1. Plasmid Construction

Plasmids pAGM4723-AflC250 and pAGM4723-eGFP350 (Figure S1) were constructedusing modular cloning with the Golden Gate gene assembly [36,37]. Briefly, sense andantisense strands of AflC were PCR-amplified from the cDNA of this gene with primerspairs AflC-F/AflC-R and AflCr-F/AflCr-R, respectively (Table S1). The pKanR promoterand rrnB-T1 terminator were obtained from plasmid pPROBE-gfp[tagless] [38] by PCRamplification with primer pairs Ppntii-F/Ppntii-R and rrnB_T1-term-F/rrnB_T1-term-R, respectively (Table S1). Sense and antisense strands of AflC were each cloned into apICH41331 (Addgene MoClo Toolkit, Watertown, MA, USA) [36,37], in addition to pKanR

and rrnB-T1, by the mixture of BpiI (Thermo Fisher Scientific, Waltham, MA, USA) and T4DNA ligase (New England Biolabs, Ipswich, MA, USA). The assemblies were done in a waythat pKanR and rrnB-T1 flanked the AflC250 fragments. Subsequently, the promoter-senseAflC250-terminator and promoter-antisense AflC250-terminator assemblies were clonedinto pICH47732 and pICH47742 (Addgene MoClo Toolkit, Watertown, MA, USA) [36,37],respectively, using BsaI instead of BpiI in the mixture. The endolysin R gene containingthe pbla promoter and rrnB-T1 terminator was PCR-amplified from plasmid pAD-LyseR(Addgene Plasmid #99244) [35] with the primer pair EndoR_F/EndoR_R (Table S1) andcloned into pICH47751 (Addgene MoClo Toolkit, Watertown, MA, USA) [36,37] by BsaIand T4 DNA ligase. Finally, the AflC250 dsRNA synthesis cassette and endolysin R genewere assembled and cloned into pAGM4723 (Addgene MoClo Toolkit, Watertown, MA,USA) [36,37], using BpiI. For cloning pAGM4723-eGFP350, sense and antisense strands ofeGFP were PCR-amplified from pICH41531 with primer pairs EGFP342-F/EGFP342-R andEGFP342-F-reversed/EGFP342-R-reversed, respectively (Table S1). pAGM4723-eGFP350was constructed by using the procedure described above.

Plasmids pAGM4723-BcSAS1250 and pAGM4723-eGFP250 (Figure S1) were constructedby adding a T7 promoter (5′–TAATACGACTCACTATAG–3′) in the forward and reverseprimers in opposite directions. BcSAS1250 was PCR-amplified from the cDNA of B. cinereawith the primer pair BcSAS1_T7_F/BcSAS1_T7_R (Table S1) and cloned into pICH47732using BsaI and T4 DNA ligase in the reaction of the Golden Gate assembly. The BcSAS1250dsRNA synthesis cassette and endolysin R gene were assembled into pAGM4723 withBpiI and T4 DNA ligase. EGFP250 was amplified from pAGM4723-eGFP350 with primerpair G250_T7_F/G250_T7_R (Table S1) and subsequently following the same proceduredescribed above to clone it into pAGM4723.

2.2. Bacterial and Fungal Strains Used in This Study, Their Growth Conditions, and InVitro Bioassays

The bacterial delivery system was based on the RNaseIII-deficient E. coli strainHT115(DE3). The genotype of HT115 is E. coli [F-, mcrA, mcrB, IN(rrnD-rrnE)1, rnc14::Tn10(DE3 lysogen: lacUV5 promoter-T7 polymerase] [29]. This strain originates from theW3110 strain, a commonly used wild-type of the non-pathogenic E. coli K-12 strain.The HT115(DE3) strain was selected for its ability to produce stable dsRNA and to expressgenes placed under the control of promoters recognized by the IPTG-inducible T7 RNApolymerase. The fungal strains analyzed were A. flavus NRRL3357 and B. cinerea B05.10.Fungal cultures were established from frozen mycelia stored in 25% glycerol (v/v) at−80 ◦C and maintained on potato dextrose agar (Difco Laboratories, Detroit, NI, USA)(PDA) plates.

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E. coli HT115(DE3) was incubated in Luria-Bertani (LB) liquid medium [10 g BactoTM

Tryptone (Difco Laboratories, Detroit, NI, USA), 10 g NaCl, 5 g BactoTM Yeast Extract(Difco Laboratories, Detroit, NI, USA), 1 L H2O, adjusted to pH 7.0] supplied with tetra-cycline (10 µg/mL) and kanamycin (50 µg/mL) at 37 ◦C at 300 rpm. The bacterialtransformants HT115/AflCCon, HT115/eGFPCon, HT115/BcSAS1Ind and HT115/eGFPIndwere generated by the heat shock method in the presence of the plasmids pAGM4723-AflC250, pAGM4723-eGFP350, pAGM4723-BcSAS1250, and pAGM4723-eGFP250, respectively.Transformants were always grown in the presence of kanamycin (50 µg/mL) and whencultures reached an optical density at 600 nm (OD600) of approximately 0.8, then isopropyl-β-D-1-thiogalactopyranoside (IPTG) (Sigma-Aldrich, St. Louis, MO, USA) was added at1.0 mM final concentration to the cultures of HT115/BcSAS1Ind and HT115/eGFPInd toinduce dsRNA production when needed. Bacterial concentration was measured with theSpectronic Genesys 6 spectrophotometer (Thermo Electron Scientific Instruments LLC,Madison, WI, USA) as optical density at 600 nm (OD600). Bacterial cultures were incubateduntil they reached the stationary phase, which was experimentally determined to be atan OD600 of 1.2–1.3 (Figure S2). The volumes corresponding to a final OD600 of 1, 0.5,0.1, and 0.01 in 10 mL cultures were concentrated by centrifugation (3000× g, 5 min) andre-suspended in molecular grade water (G-Biosciences, St. Louis, MO, USA) to a finalvolume of 0.5 mL. To induce autolysis, the bacterial cells were subjected to a freeze-thawcycle consisting of 15–20 min at −80 ◦C, followed by thawing for 60–90 min at 37 ◦C andsubsequently treating the cell suspensions with the antibiotics ampicillin (100 µg/mL finalconcentration) and streptomycin (50 µg/mL) for 1 h, to assure that no living cells werepresent. Growth curves for the HT115/AflCCon, HT115/eGFPCon, HT115/BcSAS1Ind, andHT115/eGFPInd strains as well as the HT115(DE3) strain (Figure S2), were calculated bymeasuring the OD600 with the Genesys 6 spectrophotometer (Thermospectronic) everyhour up to 9 h of growth, and then at 13 h and 24 h. All strains were grown in LB mediumsupplemented with tetracycline (10 µg/mL) and kanamycin (50 µg/mL). Kanamycinwas not added to the HT115(DE3) culture, whereas when reaching OD600 ≈ 0.8, strainsHT115/BcSAS1Ind and HT115/eGFPInd were induced with 1.0 mM IPTG. In each of thethree experiments performed, the cultures were grown at 37 ◦C in 20 mL volumes with250 rpm of shaking, with three replicates per strain.

To obtain fungal spores, A. flavus NRRL3357 was grown on PDA medium with con-tinuous light at 37 ◦C for 3–5 days (non-aflatoxin production conditions), while B. cinereawas grown on PDA medium at 25 ◦C for 10–14 days. Spores were collected by scrappingmycelia with 2 mL of milli-Q water and filtrating them through two layers of cheesecloth.Spore concentration was measured with a hemocytometer. The spore suspension wassubsequently added to 5 mL of YES liquid medium [2% BactoTM Yeast Extract (Difco Labo-ratories, Detroit, NI, USA), 6% sucrose, pH 5.8] to a final concentration of 104 spores/mL.The spore suspension was then incubated at 37 ◦C (A. flavus) or 25 ◦C (B. cinerea), 250 rpmwith continuous light. Once fungal spores had germinated (checked by microscopy at6–8 h), the bacterial treatments were added to the fungal culture flasks. These bacterialtreatments were either the living bacterial cells or their whole-cell autolysates produced in10 mL and concentrated in 0.5 mL, as described above. The co-incubation conditions forboth fungi were 25 ◦C, 250 rpm, in darkness. Fungal samples were harvested 12 h, 24 h,and 36 h after the bacterial treatment applications.

B. cinerea dry weight was measured after filtering the fungal culture with a vacuum fil-tration system using a 0.45 µm pore size nitrocellulose membrane (Thermo Fisher Scientific,Waltham, MA, USA). Once samples were completely dry, they were fully removed fromthe membrane with the help of a spatula and measured using a precision mg scale (MettlerToledo, ML203E/03). Culture filtering by vacuum filtration, performed as described above,was also used to harvest the supernatant of A. flavus cultures, of which 1 mL was usedto measure total content of aflatoxins by Enzyme-Linked Immunosorbent Assay (ELISA).Dried fungal mycelium was frozen in liquid nitrogen and stored at −80 ◦C before totalRNA extraction with the TRizolTM reagent (Invitrogen, Carlsbad, CA, USA).

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2.3. Total RNA Extractions and RT-qPCR

Total RNA was extracted from A. flavus, B. cinerea, the inoculated plant tissues, andthe bacterial strains using the TRIzolTM reagent (Invitrogen, Carlsbad, CA, USA), accord-ing to the manufacturer’s instructions. Briefly, 1 mL of TRIzolTM reagent (Invitrogen,Carlsbad, CA, USA) was added to 50–100 mg of ground plant tissue, fungal, or bacterialcells, and incubated for 5 min at room temperature. Then, 0.2 mL of chloroform wasadded to the samples and incubated for 3 min at room temperature, followed by cen-trifugation for 15 min at 12,000× g at 4 ◦C. The nucleic acids in the supernatant wereprecipitated by adding 500 µL of isopropyl alcohol (1:1) and centrifuging at 12,000× g at4 ◦C for 10 min. Pellets were washed with 75% ethanol/water (v/v), re-suspended withDEPC-treated water (Invitrogen, Carlsbad, CA, USA), and treated with DNaseI accord-ing to the manufacturer’s instructions (Thermo Fisher Scientific, Waltham, MA, USA).Following total RNA extraction, samples were tested for integrity in 1.2% agarose gels andquantified using a Nanodrop Spectrophotometer (Nanodrop 2000, Thermo Fisher Scientific,Waltham, MA, USA). The absence of DNA contamination was examined in the samplesby the lack of PCR amplification of the glyceraldehyde 3-phosphate dehydrogenase gene(GADPH) gene in A. flavus and of the BcActin gene in B. cinerea. cDNA was synthesizedfrom purified total RNA using iScriptTM cDNA Synthesis Kit (Bio-Rad Laboratories, Her-cules, CA, USA), following the manufacturer’s instructions. The SYBR Green kit SsoFastTM

EvaGreen® (Bio-Rad Laboratories, Hercules, CA, USA) was used to carry out RT-qPCRreactions in a CFX96 Touch thermal cycler (Bio-Rad laboratories, Hercules, CA, USA)equipped with the CFX MaestroTM software. The thermal profile was as recommended bythe manufacturer (40 cycles of 98 ◦C for 3 s and 60 ◦C for 5 s), testing single amplificationby a specific peak in the dissociation curve (0.5 ◦C increments every 10 s within a rangeof 65–95 ◦C). The GADPH gene, amplified with the primer pair qGADPH-F/qGADPH-R(Table S1), and the BcActin gene, amplified with the primer pair qBcActin-F/qBcActin-R(Table S1), were used as endogenous reference genes for A. flavus and B. cinerea, respec-tively. The primer pair qAflC-F/qAflC-R (Table S1) was used for AflC amplification, whileprimer pair qBcSAS1-F/qBcSAS1-R (Table S1) was used for BcSAS1. The AflC and BcSAS1sequences of the amplicons generated by RT-qPCR do not match with the dsRNA sequenceused in the treatments, to avoid artifacts. The relative expression level of each gene wascalculated by the 2−∆∆Ct method [39].

2.4. dsRNA Isolation from E. coli HT115(DE3)

Total RNA was extracted from E. coli HT115(DE3) cultures using the TRIzolTM reagent(Invitrogen, Carlsbad, CA, USA) according to the manufacturer’s instructions and asdescribed above. Then, dsRNAs were isolated from total RNA extraction, using oneof the following two methods. In the first method, 20 µg of total RNA was treated at37 ◦C for 45 min with 3 µL of DNaseI (Thermo Fisher Scientific, Waltham, MA, USA)and 1 µL of RNase A (Thermo Fisher Scientific, Waltham, MA, USA) in a 50 µL reactionvolume containing 0.35 mM NaCl and 5 µL of 10 × DNase I buffer. dsRNAs were furtherpurified with the RNA Clean & Concentrator Kit (ZYMO Research, Irvine, CA, USA)or the mirVanaTM miRNA isolation kit (Thermo Fisher Scientific, Waltham, MA, USA),following the specifications for <200 nucleotides RNA removal. In the second method,the MEGAscriptTM T7 Transcription Kit (Thermo Fisher Scientific, Waltham, MA, USA)was used from the step of nuclease digestion to remove DNA and ssRNAs from the filtercartridge provided with the kit, followed by purification of dsRNAs from the filter cartridge,which is able to remove free nucleotides and <200 nucleotides nucleic acid degradationproducts. Finally, dsRNAs were visualized by electrophoresis and their concentrationwas determined with a Nanodrop Spectrophotometer (Thermo Fisher Scientific, Waltham,MA, USA).

To examine dsRNA stability over time in the bacterial autolysates and in living bacte-rial cultures, strain HT115/BcSAS1Ind was grown to the stationary phase(OD600 ≈ 1.2–1.3) in LB medium supplemented with tetracycline (10 µg/mL) and kanamycin

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(50 µg/mL). When the culture had reached an OD600 ≈ 0.8, IPTG was added to a finalconcentration of 1.0 mM. The culture was subsequently diluted to an OD600 of 1.0 in 10 mLvolumes. After that, the 10 mL cultures were pelleted by centrifugation at 3000× g for 5 minand re-suspended in 0.5 mL of molecular grade water (G-Biosciences, St. Louis, MO, USA).The bacterial autolysates were produced as described above. Living bacterial cells wereproduced without the freeze-thaw cycle and the antibiotic treatment. Subsequently, theautolysates and the living bacterial cells were incubated at room temperature (≈ 25 ◦C) for0 h, 1 h, 6 h, 24 h, 36 h, and 72 h, after which the debris was removed by centrifugation(21,000× g, 30 min, at 4 ◦C) and total nucleic acids were extracted from the supernatantswith the TRIzolTM method as described above. The nucleic acids were then treated withRNaseA and DNase I, and cleaned with the RNA Clean & Concentrator™ Kit, as de-scribed above. Finally, aliquots of 0.5 µg of RNA were loaded onto 2% agarose gelsfor electrophoresis.

2.5. Bioassays

Fresh maize seeds were surface sterilized as described previously [40]. Briefly, theentire seeds were treated with a 75% ethanol/water (v/v) solution for 30 s at room tem-perature, then rinsed with a milli-Q autoclaved water. Next, they were treated with a 2%hypochlorite solution for 5 min at room temperature and rinsed at least five times withmilli-Q autoclaved water. Aliquots of 2 µL of a 50 × 106 spore/mL suspension (milli-Qautoclaved water) of A. flavus were placed on a maize embryo and were incubated at 37 ◦Cwith light for 2 days (non-aflatoxin production conditions). After this time period, 10 µLof living or lysed bacteria cells obtained from the OD600 1.0 culture were placed at thesame point of fungal inoculation and incubated for one more day at aflatoxin-productionconditions (darkness at 25 ◦C). Humidity was maintained to over 80% by placing themaize seeds into boxes layered with sterile Whatman filter paper 1/5, which was hydratedwith 1 mL of sterile water every 24 h. Subsequently, the maize embryos with the fungalmycelium were harvested for further analysis with RT-qPCR (40 cycles of 98 ◦C for 3 s and60 ◦C for 5 s) and ELISA.

Nicotiana benthamiana leaves were inoculated with B. cinerea, as described previ-ously [41]. Briefly, B. cinerea spores were collected from PDA plates incubated for10–14 days, as described above. Spores were purified and re-suspended at a concentrationof 104 spores/mL in grape juice (1/2 strength). A drop of 4 µL of this spore suspension wasinoculated onto both sides of detached 4–6 week-old N. benthamiana leaves, which wereplaced onto 1.5% (m/v) phytoagar (Thomas Scientific, Swedesboror, NJ, USA) mediumtrays. After 6 h, 10 µL of living or lysed bacteria cells obtained from the OD600 1.0 culturewere placed at the same point of fungal inoculation, on the left side of the leaf blade forHT115/eGFPInd and the right side of the leaf blade for HT115/BcSAS1Ind. Suspensions ofliving bacteria cells were applied with IPTG at a final concentration of 0.1 mM. InoculatedN. benthamiana leaves were incubated for 3 days. The relative lesion size between treatmentsin each leaf was determined by measuring the necrotic area using the ImageJ software.Then, they were collected for RT-qPCR analysis (40 cycles of 98 ◦C for 3 s and 60 ◦C for 5 s).

2.6. Analysis of Total Aflatoxins by ELISA

The total concentration of aflatoxins on E. coli-A. flavus co-cultures and maize seedsinfected with A. flavus was analyzed by ELISA. The AgraQuant® Total Aflatoxin Test Kit(4–40 ppb) (Romer labs®, Tulln, Austria) was used for the ELISA analysis according to themanufacturer’s instructions. Briefly, aflatoxins were extracted with 70% (v/v) methanolfrom 1 mL of fungal culture supernatants or grinded inoculated maize seeds. Because theconcentration of aflatoxins in the samples was unknown and it could fall outside of therange of the solutions of known aflatoxin concentration provided by the manufacturer(i.e., the standards), serial dilutions of the experimental samples were prepared (1:0, 1:10,1:100, and 1:1000) in 70% (v/v) methanol, so that one of the serial dilutions would fallwithin the aflatoxin concentration range covered by the standards provided with the

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manufacture’s kit (0, 4, 10, 20, 30 and 40 ppb). The OD of each sample dilution at 450nm (OD450) was measured at the end of the ELISA reactions and the sample dilutionwhose OD450 measurement was in the range of the manufacturer’s standards was selected.Subsequently, aliquots of 100 µL of the selected diluted samples and the standards in du-plicate were added to 200 µL of the conjugate. After mixing, 100 µL were transferred toantibody-coated wells and incubated for 15 min at room temperature. The unbound conju-gate was removed with deionized water during the washing step (5×). Once wells weredry, 100 µL of substrate were added into each well, incubated for 5 min, and then 100 µL ofthe stop solution were added. Microtiter plates were measured in an ELISA reader (TristarLB941, Berthold Technologies, Bad Wildbad, Germany) at 450 nm. The concentration ofaflatoxins in the samples was calculated using the formula of the calibration curve fromthe standards and multiplied by the specific dilution factor of the sample.

2.7. Statistical Analyses

Results represent the mean and the standard deviations of at least three biologicalreplicates, which are indicated by error bars. Significant differences were determined usingan unpaired t-test to compare the means between two groups, and a one-way ANOVAfollowed by Tukey’s Honestly Significant Difference (HSD) test to compare the meansbetween three or more groups. The one-sample t-test was used to test differences betweenthe relative aflatoxins production or fungal lesion size of each biological treatment withrespect to the control treatment (designated as 1). o p < 0.10; * p < 0.05; ** p < 0.01,*** p < 0.001. Statistical analyses were performed using Statistix 10.0 software package(Analytical Software, Tallahassee, FL, USA).

3. Results3.1. Bacteria Can Be Genetically Engineered to Produce dsRNAs against Target Genes in Fungi

For our studies, we have utilized the well-characterized E. coli HT115(DE3) strain,which lacks the rnc allele that encodes for the dsRNA-specific endonuclease RNaseIII, and isthus capable of synthesizing and preserving dsRNAs. The strain is also lysogenic for λDE3,thus enabling the transcription of genes placed under the control of the bacteriophage T7promoter [29]. HT115(DE3) was genetically engineered to express dsRNAs against AflCin A. flavus [42] and BcSAS1 in B. cinerea [43]. These two pathogens were selected due totheir importance as a producer of aflatoxins and a plant pathogen with a broad host range,respectively [32,34]. In addition, silencing following uptake of dsRNAs by their myceliahas been demonstrated for both fungi [12,13,15,24,40]. Likewise, AflC and BcSAS1 wereselected as gene targets for RNAi because of their critical role in production of aflatoxinsin A. flavus and virulence in B. cinerea, respectively. Specifically, AflC encodes a Type Ipolyketide synthase (pksA) needed for the synthesis of the backbone structure of aflatoxinsin A. flavus, whereas BcSAS1 encodes for a member of the Rab family of small GTPases thatis required for mycelial development and virulence in B. cinerea [42,43].

To further explore the versatility of the system, dsRNAs against AflC and BcSAS1were produced in HT115(DE3) using two plasmid vectors that allowed the productionof dsRNAs via different transcriptional configurations, i.e., (i) through the constitutivein trans (i.e., from separate plasmid loci) transcription of the sense and antisense strandsof the selected target DNA template (Figure 1A) and (ii) through the IPTG-inducible incis (i.e., from the same plasmid locus) bi-directional transcription of the DNA template(Figure 1B). Specifically, in the first configuration that was used to produce dsRNAs againstAflC, a 250 bp cDNA fragment was cloned in opposite directions between the native pro-moter of the kanamycin resistance gene (pKanR) and the transcriptional terminator T1of the E. coli rrnB gene (rrnB-T1). A plasmid construct similar to this one but in whichthe AflC fragment was replaced by a 350 bp cDNA fragment of the enhanced green flu-orescent protein (eGFP) was also prepared and used in control applications (Figure 1Aand Figure S1A). In the second configuration that was used to produce dsRNAs againstBcSAS1, two T7 promoters (pT7) were placed in an inverted orientation at the 5′ and 3′-end

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of a 250 bp cDNA fragment of the gene (Figure 1B). As for AflC, a plasmid construct inwhich the BcSAS1 fragment was replaced by a 250 bp eGFP cDNA fragment was alsoprepared and used as control (Figure 1B and Figure S1B). Following transformation intoHT115(DE3) of the plasmid constructs and transcription, the first configuration producesin trans complementary sense and antisense ssRNA molecules from the DNA templates,which subsequently hybridize to form dsRNAs (Figure 1C). By contrast, in the secondconfiguration sense/antisense transcript pairs are spontaneously formed in cis, thus cre-ating dsRNAs (Figure 1C). Moreover, while in the first configuration the transcriptionof the target sequences is controlled by the constitutive pKanR promoter, in the secondconfiguration it is dependent on the induction of the T7 gene 1 in HT115(DE3) by IPTGand subsequent activation of the T7 promoter by the T7 RNA polymerase.

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AflC, a 250 bp cDNA fragment was cloned in opposite directions between the native pro-moter of the kanamycin resistance gene (pKanR) and the transcriptional terminator T1 of the E. coli rrnB gene (rrnB-T1). A plasmid construct similar to this one but in which the AflC fragment was replaced by a 350 bp cDNA fragment of the enhanced green fluorescent protein (eGFP) was also prepared and used in control applications (Figure 1A and Figure S1A). In the second configuration that was used to produce dsRNAs against BcSAS1, two T7 promoters (pT7) were placed in an inverted orientation at the 5′ and 3′-end of a 250 bp cDNA fragment of the gene (Figure 1B). As for AflC, a plasmid construct in which the BcSAS1 fragment was replaced by a 250 bp eGFP cDNA fragment was also prepared and used as control (Figure 1B and Figure S1B). Following transformation into HT115(DE3) of the plasmid constructs and transcription, the first configuration produces in trans com-plementary sense and antisense ssRNA molecules from the DNA templates, which sub-sequently hybridize to form dsRNAs (Figure 1C). By contrast, in the second configuration sense/antisense transcript pairs are spontaneously formed in cis, thus creating dsRNAs (Figure 1C). Moreover, while in the first configuration the transcription of the target se-quences is controlled by the constitutive pKanR promoter, in the second configuration it is dependent on the induction of the T7 gene 1 in HT115(DE3) by IPTG and subsequent ac-tivation of the T7 promoter by the T7 RNA polymerase.

As both trans-kingdom transfer and environmental uptake of RNAi molecules by mycelial cells have been demonstrated to induce RNAi in fungi, in order to combine the benefits of the two approaches, we further genetically engineered the HT115(DE3) strain to allow the release of the dsRNA cargo through a programmable cellular autolysis step. For this purpose, a phage lambda endolysin gene (λR), placed under the control of the weakly constitutive pbla promoter of the β-lactamase gene and the rrnB-T1 terminator was cloned in the two plasmid vectors used for the production of the dsRNAs (Figure 1A,B and Figure S1A,B). When the inner bacterial cell membrane is intact, the endolysin protein accumulates in the cell without significantly impacting cell growth. However, a simple freeze-thaw cycle opens the inner membrane, thus enabling the endolysin to disrupt the bacterial cell wall and trigger autolysis [35]. Notably, although cell lysis results in the re-lease of several lytic enzymes, the autolysis of the bacterial cell did not trigger a parallel degradation of the dsRNAs, which remained stable in the cell lysates up to 24 h, and par-tially stable up to 72 h of incubation at 25 °C. If the cells are not lysed, then dsRNAs remain stable in the living cells for at least for 72 h (Figure 1D). This indicates that both living bacteria and their lysates can be used to deliver dsRNAs against fungi.

Figure 1. Bacteria can be genetically engineered to produce dsRNAs against target genes in fungi. (A, B) Schematic repre-sentation of the two transcriptional configurations used for the synthesis of dsRNAs against AflC in A. flavus and BcSAS1 Figure 1. Bacteria can be genetically engineered to produce dsRNAs against target genes in fungi. (A,B) Schematicrepresentation of the two transcriptional configurations used for the synthesis of dsRNAs against AflC in A. flavus andBcSAS1 in B. cinerea, respectively as well as for the synthesis of dsRNA against the enhanced green fluorescent protein (eGFP).The full plasmid maps are shown in Figure S1. pKanR: kanamycin resistance gene; rrnB-T1: transcriptional terminatorT1 of the rrnB gene; pT7: T7 RNA polymerase promoter; pbla: promoter of the β-lactamase gene; λR: phage lambdaendolysin gene; AflC: Type I polyketide synthase (pksA) in A. flavus; BcSAS1: Rab family member of small GTPases; eGFP:enhanced green fluorescent protein. (C) Agarose gel electrophoresis of dsRNAs extracted from the autolyzed HT115/AflCCon,HT115/BcSAS1Ind, HT115/eGFPCon, and HT115/eGFPInd strains. Production of dsRNAs in the HT115/BcSAS1Ind andHT115/eGFPInd strains requires the addition of IPTG in the culture medium of the bacteria, which was added at a finalconcentration of 1.0 mM when cultures reached an OD600 ≈ 0.8. (D) Stability over time of the dsRNAs produced by theHT115/BcSAS1Ind strain over time. Stability was examined both in living bacteria cells (left-hand side image) and theirwhole-cell autolysates (right-hand side image). An equal volume of 5 µL was loaded in the wells of the agarose gel.

As both trans-kingdom transfer and environmental uptake of RNAi molecules bymycelial cells have been demonstrated to induce RNAi in fungi, in order to combine thebenefits of the two approaches, we further genetically engineered the HT115(DE3) strainto allow the release of the dsRNA cargo through a programmable cellular autolysis step.For this purpose, a phage lambda endolysin gene (λR), placed under the control of theweakly constitutive pbla promoter of the β-lactamase gene and the rrnB-T1 terminator wascloned in the two plasmid vectors used for the production of the dsRNAs (Figure 1A,Band Figure S1A,B). When the inner bacterial cell membrane is intact, the endolysin proteinaccumulates in the cell without significantly impacting cell growth. However, a simplefreeze-thaw cycle opens the inner membrane, thus enabling the endolysin to disrupt thebacterial cell wall and trigger autolysis [35]. Notably, although cell lysis results in the

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release of several lytic enzymes, the autolysis of the bacterial cell did not trigger a paralleldegradation of the dsRNAs, which remained stable in the cell lysates up to 24 h, andpartially stable up to 72 h of incubation at 25 ◦C. If the cells are not lysed, then dsRNAsremain stable in the living cells for at least for 72 h (Figure 1D). This indicates that bothliving bacteria and their lysates can be used to deliver dsRNAs against fungi.

3.2. Bacterially-Produced dsRNAs Induce RNAi in Fungi in a BacteriaConcentration-Dependent Manner

We initially sought to determine whether the HT115(DE3) bacterial strains thatwere genetically engineered to produce dsRNAs against AflC [HT115(DE3)(pAGM4723-AflC250)], hereafter abbreviated as HT115/AflCCon, or BcSAS1 [HT115(DE3)(pAGM4723-BcSAS1250)], hereafter abbreviated as HT115/BcSAS1Ind, were able to induce silencingof the targeted transcripts in A. flavus and B. cinerea, respectively. Two approaches fordelivering the bacterially-produced dsRNAs against A. flavus and B. cinerea were utilized,i.e., trans-kingdom mediated transfer, facilitated by co-incubating living bacteria withfungal germlings (hereafter referred to as Treatment A), and environmental uptake frombacterial whole-cell autolysates (hereafter referred to as Treatment B).

To examine whether the physical interaction between fungi and bacteria can facilitatethe trans-kingdom transfer of the bacterially-produced dsRNAs to fungi (Treatment A),10 mL bacterial cultures with a final OD600 of 1, 0.5, 0.1, and 0.01 that were prepared fromserially-diluted cultures at stationary phase (OD600 ≈ 1.2–1.3) (Figure S2), were concen-trated to a volume of 0.5 mL and mixed with 5 mL of fungal germlings (104 germlings/mL)in liquid cultures. Assessment of RNAi-induced gene silencing at 12 h of co-incubation be-tween A. flavus and the HT115/AflCCon strain, or between B. cinerea and the HT115/BcSAS1Indstrain (hereafter referred to as test applications), showed a significant reduction in AflC andBcSAS1 transcript levels compared with the expression levels of these two genes whenthe two fungi were co-incubated with the HT115/eGFPCon and HT115/eGFPInd strains,respectively (hereafter referred to as control applications) (Figure 2A,B, and Table S2).Overall, when germlings of A. flavus or B. cinerea were co-incubated with the bacterialconcentrates obtained from the OD600 1.0, 0.5, and 0.1 cultures, a 21.7-fold, a 5.5-fold, and a3.2-fold reduction in AflC expression, and a 23.3-fold, a 15.9-fold, and a 2.7-fold reductionin BcSAS1 expression were observed in the test applications relative to the control ones,respectively (Figure 2A,B and Table S2). These reductions in AflC and BcSAS1 transcriptlevels between the test and control applications were significant at α = 0.05 (Table S2) andpositively correlated with the OD600 of the bacterial cultures (AflC: r = 0.74, p = 0.0061;BcSAS1: r = 0.73, p = 0.007). Moreover, the reduction in target gene expression was pre-sumably caused by the bacterially-produced dsRNAs alone, as the expression levels ofAflC and BcSAS1 did not statistically differ among the control applications, or betweenthe control applications and the axenic culture of A. flavus and B. cinerea (Figure S3A,B,and Tables S3 and S4). Taken together, the above results indicate that the two HT115(DE3)strains that were genetically engineered to produce dsRNAs against AflC or BcSAS1 arecapable of inducing RNAi to A. flavus and B. cinerea in in vitro co-cultures of the twoorganisms, in a bacteria concentration-dependent manner.

As for Treatment A, a bacteria concentration-dependent response in the silencinglevels of AflC and BcSAS1 (AflC: r = 0.80, p = 0.0017; BcSAS1: r = 0.77, p = 0.0037) wasalso observed when germlings of A. flavus and B. cinerea were incubated for 12 h underthe same experimental conditions with the whole-cell autolysates of HT115/AflCCon andHT115/BcSAS1Ind, respectively (Treatment B) (Figure 2A,B and Table S2). Specifically, a sig-nificant (p < 0.05) reduction in the expression levels of AflC in the test relative to the controlapplications was obtained when A. flavus was incubated with lysates of HT115/AflCCon col-lected from the OD600 1.0 and 0.5 cultures but not with the lysates collected from the OD6000.1 and 0.01 cultures (Figure 2A and Table S2). On a quantitative basis, subtle changesin AflC expression between test and control applications were observed in Treatment Bcompared with Treatment A for all bacterial densities tested (Table S2), suggesting thatenvironmental uptake of dsRNAs from bacterial whole-cell autolysates was less efficient in

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eliciting RNAi in A. flavus compared with applications of living bacteria. When consideringBcSAS1, then a significant (p < 0.05) reduction in its expression level was observed whengermlings of B. cinerea were co-incubated with lysates of HT115/BcSAS1Ind collected fromall four serially diluted cultures (i.e., cultures with an OD600 of 1, 0.5, 0.1, 0.01) (Figure 2Band Table S2). Moreover, fold-changes in BcSAS1 transcript levels between test and controlapplications were lower in Treatment B compared with Treatment A for the OD600 1.0and 0.5 cultures, but higher for the OD600 0.1 and 0.01 cultures (Figure 2B and Table S2).This suggests that higher densities of cell lysates could be interfering with the absorptionof dsRNAs by the hyphae, thus making it appear that living bacteria are more potent ineliciting silencing. Finally, the inclusion of just the bacterial lysates in the liquid cultureshad no effect on the expression of AflC or BcSAS1, as their transcript levels were essentiallythe same among control applications (Figure S3A,B and Tables S3 and S4).

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Figure 2. Bacterially-produced dsRNAs induce RNAi in fungi in a bacteria concentration-dependent manner. (A,B) Ex-pression levels of AflC from A. flavus (Panel A) and BcSAS1 from B. cinerea (Panel B), in in vitro cultures of the fungi with living (Treatment A) or lysed (Treatment B) E. coli HT115(DE3) bacteria producing dsRNAs against AflC (HT115/AflCCon) or BcSAS1 (HT115/BcSAS1Ind), respectively (test applications; blue bars). Expression levels are shown relative to the ex-pression of AflC and BcSAS1 when A. flavus and B. cinerea are co-incubated with the control HT115(DE3) bacteria produc-ing dsRNAs against eGFP (HT115/eGFPCon and HT115/eGFPInd, respectively) (control applications; grey bars set to 1.0). Liv-ing bacteria or their whole-cell autolysates, obtained from the OD600 0.01, 0.1, 0.5, and 1.0 cultures were added to the fungal cultures, and the expression of AflC or BcSAS1 was measured at 12 h of co-incubation. For comparison, AflC and BcSAS1 silencing levels achieved by the addition in the in vitro cultures of A. flavus or B. cinerea of 15 μg (12.5 nM) of purified dsRNAs against each gene, respectively were also evaluated. Significant differences between control and test applications were determined by a one-way ANOVA followed by a Tukey’s HSD test, and levels of significance are indicated above the bars by underlined star symbols (i.e., * p < 0.05, ** p < 0.01). Differences in the gene silencing efficacy of AflC and BcSAS1 among the test applications were tested by ANOVA and Tukey’s HSD test, and significantly different groups (p < 0.05) are indicated by letters above the bars. Error bars in the figure indicate standard deviation (SD) obtained from three bio-logical replicates.

As for Treatment A, a bacteria concentration-dependent response in the silencing lev-els of AflC and BcSAS1 (AflC: r = 0.80, p = 0.0017; BcSAS1: r = 0.77, p = 0.0037) was also observed when germlings of A. flavus and B. cinerea were incubated for 12 h under the same experimental conditions with the whole-cell autolysates of HT115/AflCCon and HT115/BcSAS1Ind, respectively (Treatment B) (Figure 2A,B and Table S2). Specifically, a significant (p < 0.05) reduction in the expression levels of AflC in the test relative to the control applications was obtained when A. flavus was incubated with lysates of HT115/AflCCon collected from the OD600 1.0 and 0.5 cultures but not with the lysates col-lected from the OD600 0.1 and 0.01 cultures (Figure 2A and Table S2). On a quantitative basis, subtle changes in AflC expression between test and control applications were ob-served in Treatment B compared with Treatment A for all bacterial densities tested (Table

Figure 2. Bacterially-produced dsRNAs induce RNAi in fungi in a bacteria concentration-dependent manner.(A,B) Expression levels of AflC from A. flavus (Panel A) and BcSAS1 from B. cinerea (Panel B), in in vitro cultures ofthe fungi with living (Treatment A) or lysed (Treatment B) E. coli HT115(DE3) bacteria producing dsRNAs against AflC(HT115/AflCCon) or BcSAS1 (HT115/BcSAS1Ind), respectively (test applications; blue bars). Expression levels are shownrelative to the expression of AflC and BcSAS1 when A. flavus and B. cinerea are co-incubated with the control HT115(DE3)bacteria producing dsRNAs against eGFP (HT115/eGFPCon and HT115/eGFPInd, respectively) (control applications; greybars set to 1.0). Living bacteria or their whole-cell autolysates, obtained from the OD600 0.01, 0.1, 0.5, and 1.0 cultures wereadded to the fungal cultures, and the expression of AflC or BcSAS1 was measured at 12 h of co-incubation. For comparison,AflC and BcSAS1 silencing levels achieved by the addition in the in vitro cultures of A. flavus or B. cinerea of 15 µg (12.5 nM)of purified dsRNAs against each gene, respectively were also evaluated. Significant differences between control and testapplications were determined by a one-way ANOVA followed by a Tukey’s HSD test, and levels of significance are indicatedabove the bars by underlined star symbols (i.e., * p < 0.05, ** p < 0.01). Differences in the gene silencing efficacy of AflCand BcSAS1 among the test applications were tested by ANOVA and Tukey’s HSD test, and significantly different groups(p < 0.05) are indicated by letters above the bars. Error bars in the figure indicate standard deviation (SD) obtained fromthree biological replicates.

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In order to appreciate the silencing levels achieved against AflC and BcSAS1 in Treat-ments A and B, an additional application of purified dsRNAs was evaluated, consistingof 15 µg (12.5 nM) of dsRNAs against AflC or BcSAS1 applied to 5 mL of in vitro cultures(104 germlings/mL) of A. flavus or B. cinerea, respectively. Compared with the expressionlevels of these two genes in control axenic cultures of A. flavus or B. cinerea, the applicationof the purified dsRNAs led to a 2.9-fold reduction in AflC transcript levels (Figure 2A)and an 8.9-fold reduction in BcSAS1 transcript levels (Figure 2B). These changes in geneexpression are intermediate between those obtained in Treatments A and B at a bacterialOD600 of 0.1 and 0.5. Thus, collectively our results indicate that the bacterial-based platformto produce dsRNAs can be effectively utilized to silence fungal genes, achieving resultsthat are on par with those obtained with applications of naked dsRNAs.

3.3. Increased Contact Times between Fungi and dsRNA-Producing Bacteria Have a Positive Effecton Reducing Production of Aflatoxins in A. flavus and Mycelial Growth in B. cinerea

We next assessed whether an increase in contact time between HT115/AflCCon andA. flavus or between HT115/BcSAS1Ind and B. cinerea, would have an added positive effecton the dsRNA-mediated induced silencing of AflC and BcSAS1, respectively.For this purpose, concentrates of living bacteria or of their whole-cell autolysates from theOD600 1.0 cultures were added to 5 mL cultures of A. flavus or B. cinerea, and silencing levelsof AflC and BcSAS1 were assessed after 12 h, 24 h, and 36 h of co-incubation (Table S5).All other conditions in these experiments were kept the same as in the in vitro experimentsdescribed above, including control applications.

Examination of the AflC transcript levels in just the control applications showed thatthe expression of this gene decreased over time in both Treatment A and Treatment B.The changes in AflC transcript levels from 12 h to 24 h and 36 h were significant at α = 0.05(12 h to 24 h, Treatment A and B) or α = 0.10 (12 h to 36 h, Treatment A). However, therewere not significant differences in AflC expression levels between 24 h and 36 h for neitherTreatment A nor Treatment B (Figure S4A and Table S6). When assessing AflC silencinglevels in the test relative to the control applications, then in Treatment A, a large 21.7-folddecrease in the AflC expression levels was observed at 12 h, but only a 5.2-fold and a 4.6-folddecrease were observed at 24 h and 36 h, respectively. The decreases in AflC expression at12 h and 24 h were significant at α = 0.05, whereas the decrease at 36 h was significant onlyat α = 0.1. In a similar way, in Treatment B, a 4.1-fold, a 1.7-fold, and a 2.6-fold decreasein AflC expression were seen at 12 h, 24 h, and 36 h, respectively in the test relative tothe control applications, of which only the decrease at 12 h was significant at α = 0.05(Figure 3A and Table S5). Collectively, the data suggest that although the HT115/AflCConliving bacteria or of their lysates strongly repressed AflC transcript levels during the first12 h, the silencing effect was nearly alleviated by 24 h, despite the continuous presence ofthe bacteria or their lysates in the co-cultures. However, it is also possible that the absenceof detectable silencing of AflC at 24 h and 36 h post bacterial applications is caused by thelow expression levels of this gene at these two time points that diminishes the effects of thedsRNA treatment.

Since AflC encodes for a key enzyme in the biosynthetic pathway of aflatoxins [42],we next examined whether the observed HT115/AflCCon-mediated silencing of AflC wouldinfluence their production. Quantification of aflatoxins content in co-cultures of TreatmentA and B after 12 h, 24 h, and 36 h showed that, unlike AflC transcript levels, the total amountof these mycotoxins increased over time, most likely because of the parallel increase andaccumulation in fungal biomass (Figure 3B and Table S7). However, when comparingaflatoxins content between test and control applications of either Treatment A or B, then adecrease was observed in the test applications that was more pronounced at 36 h, followedby 24 h and 12 h. Specifically, in co-cultures of Treatment A, a 64.6% and a 30.5% reductionin aflatoxins content was observed in the test relative to the control applications at 36 h and24 h of co-incubation time, respectively, whereas at 12 h, a 13.3% increase was observed.The decrease at 36 h was significant at α = 0.05, whereas at 24 h it was significant at α = 0.1and at 12 h it was not significant (Figure 3B and Table S7). In a similar way, in co-cultures of

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Treatment B, a 42.3%, 35.4%, and 0% reduction in aflatoxins content was observed in the testrelative to the control application at 36 h, 24 h, and 12 h of co-incubation time, respectively.The differences were significant at α = 0.05 at 36 h and 24 h but not significant at 12 h (Figure3B and Table S7). Overall, our results show that bacterially-produced dsRNAs against AflCcan significantly impair production of aflatoxins over time, whereas the inverse correlationseen between AflC mRNA abundance and concentration of aflatoxins, suggests that thereis likely a delay between AflC expression, and biosynthesis and secretion of aflatoxins.

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by the low expression levels of this gene at these two time points that diminishes the ef-fects of the dsRNA treatment.

Figure 3. Increased contact times between fungi and dsRNA-producing bacteria have a positive effect on reducing pro-duction of aflatoxins in A. flavus and mycelial growth in B. cinerea. (A,B) AflC expression levels over time (Panel A) and the amount of aflatoxins produced by A. flavus (Panel B) in in vitro cultures of the fungus with living (Treatment A) or lysed (Treatment B) E. coli HT115(DE3) bacteria producing dsRNAs against AflC (HT115/AflCCon; test application) or eGFP (HT115/eGFPCon; control application). Living bacteria or their whole-cell autolysates, obtained from the OD600 1.0 culture, were mixed with the fungal cultures, and AflC expression or the amount of aflatoxins in the co-cultures were measured at 12 h, 24 h, and 36 h. In Panel A, the expression levels of AflC in the test applications (blue bars) are shown relative to the control ones (grey bars, set to 1.0). Significant differences between control and test applications were determined by a one-way ANOVA followed by a Tukey’s HSD test, and levels of significance are indicated above the bars by underlined star symbols (i.e., o p < 0.1, * p < 0.05, ** p < 0.01). Differences in the gene silencing efficacy of AflC among the test applications were tested by ANOVA and a Tukey’s HSD test, and significantly different groups (p < 0.05) are indicated by letters above the bars. In Panel B, yellow bars indicate the mean relative amount of aflatoxins produced by the fungus in the test relative to control applications, with the latter set to 100% and shown as the background grey bars. The slope analysis in Panel B shows the actual amount of aflatoxins in ppb produced over time by the fungus in control (grey lines) and test applications (blue lines). Error bars in Panels A and B indicate standard deviation (SD) obtained from three biological replicates. Sig-nificant differences in the relative mean amount of aflatoxins between control and test applications were determined by one-sample t-test, while significant differences in the absolute amount of aflatoxins were determined by a one-way ANOVA followed by a Tukey’s HSD test. Levels of significance are indicated above the bars by blue star symbols (i.e., o p < 0.1, * p < 0.05, ** p < 0.01). (C,D) Similar to Panels A and B, but showing BcSAS1 expression levels (Panel C) and the amount of biomass produced by B. cinerea (Panel D) over time in in vitro cultures of the fungus with living (Treatment A) or lysed (Treatment B) E. coli HT115(DE3) bacteria producing dsRNAs against BcSAS1 (HT115/BcSAS1Ind; test application) or eGFP (HT115/eGFPInd; control application).

Since AflC encodes for a key enzyme in the biosynthetic pathway of aflatoxins [42], we next examined whether the observed HT115/AflCCon-mediated silencing of AflC would influence their production. Quantification of aflatoxins content in co-cultures of Treat-ment A and B after 12 h, 24 h, and 36 h showed that, unlike AflC transcript levels, the total amount of these mycotoxins increased over time, most likely because of the parallel in-crease and accumulation in fungal biomass (Figure 3B and Table S7). However, when comparing aflatoxins content between test and control applications of either Treatment A

Figure 3. Increased contact times between fungi and dsRNA-producing bacteria have a positive effect on reducingproduction of aflatoxins in A. flavus and mycelial growth in B. cinerea. (A,B) AflC expression levels over time (Panel A)and the amount of aflatoxins produced by A. flavus (Panel B) in in vitro cultures of the fungus with living (Treatment A) orlysed (Treatment B) E. coli HT115(DE3) bacteria producing dsRNAs against AflC (HT115/AflCCon; test application) or eGFP(HT115/eGFPCon; control application). Living bacteria or their whole-cell autolysates, obtained from the OD600 1.0 culture,were mixed with the fungal cultures, and AflC expression or the amount of aflatoxins in the co-cultures were measuredat 12 h, 24 h, and 36 h. In Panel A, the expression levels of AflC in the test applications (blue bars) are shown relative tothe control ones (grey bars, set to 1.0). Significant differences between control and test applications were determined by aone-way ANOVA followed by a Tukey’s HSD test, and levels of significance are indicated above the bars by underlined starsymbols (i.e., o p < 0.1, * p < 0.05, ** p < 0.01). Differences in the gene silencing efficacy of AflC among the test applicationswere tested by ANOVA and a Tukey’s HSD test, and significantly different groups (p < 0.05) are indicated by letters abovethe bars. In Panel B, yellow bars indicate the mean relative amount of aflatoxins produced by the fungus in the test relativeto control applications, with the latter set to 100% and shown as the background grey bars. The slope analysis in Panel Bshows the actual amount of aflatoxins in ppb produced over time by the fungus in control (grey lines) and test applications(blue lines). Error bars in Panels A and B indicate standard deviation (SD) obtained from three biological replicates.Significant differences in the relative mean amount of aflatoxins between control and test applications were determined byone-sample t-test, while significant differences in the absolute amount of aflatoxins were determined by a one-way ANOVAfollowed by a Tukey’s HSD test. Levels of significance are indicated above the bars by blue star symbols (i.e., o p < 0.1,* p < 0.05, ** p < 0.01). (C,D) Similar to Panels A and B, but showing BcSAS1 expression levels (Panel C) and the amount ofbiomass produced by B. cinerea (Panel D) over time in in vitro cultures of the fungus with living (Treatment A) or lysed(Treatment B) E. coli HT115(DE3) bacteria producing dsRNAs against BcSAS1 (HT115/BcSAS1Ind; test application) or eGFP(HT115/eGFPInd; control application).

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When examining how increased contact times between B. cinerea and living HT115/BcSAS1Ind or the lysates thereof affected BcSAS1 transcript levels and subsequent growthof the fungus, then a positive correlation was observed as well. Overall, BcSAS1 expressionlevels increased from 12 h to 24 h and 36 h in the control applications of both Treatment Aand Treatment B. However, the differences in BcSAS1 expression between 12 h, 24 h, and36 h were not significantly different from each other (Figure S4B and Table S8), indicatingthat the expression of this gene was essentially stable in the controls throughout thecourse of the experiment. When compared with the control applications, exposure ofB. cinerea to living HT115/BcSAS1Ind bacterial cells or their whole-cell autolysates resultedin a significant at α = 0.01 reduction in BcSAS1 transcript levels at 12 h and 36 h of co-incubation time. However, at 24 h, the reduction was significant only at α = 0.10 (Figure 3Cand Table S5). Specifically, in co-cultures of Treatment A, a 23.3-fold and 14.5-fold decreasein BcSAS1 transcripts levels were observed in the test applications relative to the controlones at 12 h and 36 h, respectively whereas only a 4.3-fold reduction was seen at 24 h.In a similar way, in co-cultures of Treatment B, a 20.0-fold and a 25.6-fold reduction wereobserved at 12 h and 36 h, respectively, whereas a 2.9-fold reduction was seen at 24 h(Figure 3C and Table S5). These oscillations in BcSAS1 silencing levels from 12 h to 24 h and36 h could represent an attempt by the fungal cells to partially recover from the effects ofRNAi, perhaps through the activation of a regulatory feedback mechanism that increasedBcSAS1 expression at around 24 h in an effort to compensate for the reduction in its mRNAlevels. Irrespective of whether such a negative-feedback loop is operational for BcSAS1,overall, the results indicate that increased contact times between the fungus and the dsRNA-producing bacteria can have a positive impact on silencing of target genes, although thisactivity seems to be dependent on the gene target and possibly other parameters as well.

Since BcSAS1 is involved in mycelial development in B. cinerea [43], we also exam-ined whether the observed HT115/BcSAS1Ind-mediated silencing of BcSAS1 at 12 h, 24 h,and 36 h of co-incubation time between the fungus and the living bacterial cells or theirautolysates would impact fungal growth. Indeed, a 16.0%, 27.4%, and 24.1% reductionin fungal dry weight was observed in the test relative to the control co-cultures of Treat-ment A at 12 h, 24 h, and 36 h of co-incubation time between the fungus and the bacteria,respectively (Figure 3D and Table S9). Although small, these changes were significant atα = 0.05 for 12 h and 24 h but not at 36 h (Table S9). In a similar way, a 39.2%, 61.3%, and24.1% reduction in fungal dry weight was seen at 12 h, 24 h, and 36 h, respectively in thetest co-cultures of Treatment B relative to the control ones (Figure 3D and Table S9). It isworth noticing that the oscillations observed in the reduction of fungal biomass at 12 h,24 h, and 36 h inversely mirrored the fluctuations observed in the BcSAS1 silencing levelsat these time points, suggesting that there could be a delay between inhibition of mRNAand phenotypic expression of growth impedance. This was more pronounced in TreatmentB, as fungal growth in Treatment A was partially inhibited by growth of the bacteria, thusalleviating some of the observed differences. In this respect, fungal growth as determinedby mycelial dry weight in the control co-cultures of Treatment A was reduced by 36.1%at 12 h (p = 0.068), 41.7% at 24 h (p = 0.003), and 16.9% at 36 h (p = 0.213) compared withTreatment B (Table S9). Despite these differences between Treatment A and Treatment B,overall the results demonstrate that applications of bacteria producing dsRNAs againstBcSAS1 can effectively reduce mycelial growth of B. cinerea under in vitro conditions.

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3.4. In Vivo Applications of dsRNA-Producing Bacteria Result in a Reduction of AflatoxinsProduction by A. flavus and Disease Symptoms Caused by B. cinerea

The in vitro experiments demonstrated that HT115/AflCCon and HT115/BcSAS1Indcould be used as vectors for delivering RNAi against the target genes in A. flavus andB. cinerea, and thus impede production of aflatoxins and mycelial growth, respectively.Therefore, we next examined whether such an effect could also be achieved under in vivoconditions during fungal infections of host plants. For this purpose, A. flavus and B. cinereawere spot inoculated on maize seeds and leaves of N. benthamiana, respectively and aftera period of incubation of 48 h for A. flavus and 6 h for B. cinerea, 10 µL of living bacterialconcentrates or their whole-cell autolysates from the OD600 1.0 cultures were applied tothe same fungal inoculation spots.

Examination of AflC transcript levels in A. flavus inoculated on maize seeds (Figure 4A)and exposed for 24 h at living HT115/AflCCon bacteria or their autolysates, showed a sig-nificant (α = 0.05) 4.8-fold and a 6.5-fold reduction, respectively in its expression levelsin the test relative to the control applications (Figure 4B and Table S10). Notably, AflCexpression levels did not statistically differ between control applications of Treatment Aand Treatment B (p = 0.441), indicating that silencing of AflC was caused by the dsRNAsproduced by the HT115/AflCCon bacteria. Moreover, the decrease in AflC transcript levelswas further accompanied by a reduction in production of aflatoxins, as determined by par-allel measurements of total aflatoxins content in the maize seeds (Figure 4C and Table S10).Specifically, for Treatment A, a 47.4% reduction was observed in the total amount of afla-toxins extracted from seeds treated with the HT115/AflCCon strain compared with seedstreated with the HT115/eGFPCon strain. In a similar way, for Treatment B, a 51.8% reductionin aflatoxins content was observed in seeds included in the test application relative to seedsincluded in the control one (Figure 4C and Table S10). The reduction in the total aflatoxinscontent between test and control applications clearly demonstrates that the HT115/AflCConstrain can effectively obstruct the production of these mycotoxins by A. flavus, also underin vivo conditions.

As for AflC, application of the living HT115/BcSAS1Ind cells or their whole-cell au-tolysates on N. benthamiana leaves inoculated with B. cinerea (Figure 5A) resulted in silencingof BcSAS1. Indeed, a 2.6-fold and 1.7-fold reduction in BcSAS1 expression was seen in thetest applications relative to the control ones for Treatment A and Treatment B, respectivelyat 72 h post addition of the living bacteria or their whole-cell autolysates on N. benthamianaleaves inoculated with B. cinerea (Figure 5B and Table S11). These differences in BcSAS1transcript levels between test and control applications were significant at α = 0.05, whereasBcSAS1 expression levels were the same between control applications of Treatment Aand Treatment B (p = 0.820), indicating that the reduction in its transcript levels in thetest applications was caused by the dsRNAs produced by the HT115/BcSAS1Ind strain.Moreover, silencing of BcSAS1 was phenotypically expressed as a parallel reduction in thesize of the infection lesions caused by B. cinerea on leaves of N. benthamiana (Figure 5A,C).Indeed, for Treatment A, a significant 35% reduction in lesion size between test and controlapplication was observed, whereas in Treatment B, this significant reduction was by 25%(Figure 5C and Table S11). Collectively, these results demonstrate that applications ofbacteria that produce dsRNAs against genes in target pathogens can be potentially usedfor disease control.

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Figure 4. The HT115/AflCCon strain induces RNAi in A. flavus and a reduction in production of aflatoxins under in vivoconditions. (A) A. flavus inoculations on maize kernels. Spores of the fungus were spot inoculated on maize kernels and48 h later, 10 µL of the living bacteria or of their whole-cell autolysates, obtained from the OD600 1.0 culture, were applied tothe same inoculation spot. All measurements were taken 24 h later. (B,C) AflC expression levels (Panel B) and the amountof aflatoxins produced by A. flavus in vivo on maize kernels (Panel C), following its exposure to living (Treatment A) orlysed (Treatment B) E. coli HT115(DE3) bacteria producing dsRNAs against AflC (HT115/AflCCon; test application) or eGFP(HT115/eGFPCon; control application). In Panel B, the expression level of AflC in the test applications (blue bars) is shownrelative to the control applications (grey bars, set to 1.0). In Panel C, charts on the left-hand side show the mean relativeaflatoxins content in the maize kernels in the test (yellow bars) relative to the control applications (grey bars, set to 100%)and, on the right-hand side, the actual amount of aflatoxins (in ppb) in the test and the control applications. Error bars inPanels B and C indicate standard deviations (SD) obtained from at least three biological replicates. Significant differences ingene expression and the absolute amount of aflatoxins between control and test applications were determined by a one-wayANOVA followed by a Tukey’s HSD test. Significant differences in the mean relative aflatoxins content were determined byone-sample t-test. Levels of significance are indicated above the bars by underlined star symbols (i.e., o p < 0.1, * p < 0.05,** p < 0.01).

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(Figure 5C and Table S11). Collectively, these results demonstrate that applications of bac-teria that produce dsRNAs against genes in target pathogens can be potentially used for disease control.

Figure 5. The HT115/BcSAS1Ind strain induces RNAi in B. cinerea and a reduction in fungal growth under in vivo conditions. (A) B. cinerea inoculations on N. benthamiana leaves. Spores of the fungus were spot inoculated on leaves of N. benthamiana and 6 h later, 10 μL of the living bacteria or the whole-cell autolysates, obtained from the OD600 1.0 culture, were applied to the same inoculation spots. All measurements were taken 72 h later. (B,C) BcSAS1 expression levels (Panel B) and the relative infection lesion size produced by B. cinerea in vivo on leaves of N. benthamiana (Panel C), following its exposure to living (Treatment A) or lysed (Treatment B) E. coli HT115(DE3) bacteria producing dsRNAs against BcSAS1 (HT115/BcSAS1Ind; test application) or eGFP (HT115/eGFPInd; control application). In Panels B and C, BcSAS1 expression levels and the mean relative lesion size of the infection areas in the test applications (blue bars and yellow bars, respec-tively) are relative to the control ones (grey bars, set to 1.0 and 100%, respectively). Error bars indicate standard deviations (SD) obtained from at least three biological replicates. Significant differences in gene expression between control and test applications were determined by a one-way ANOVA followed by Tukey’s HSD test, whereas significant differences in the mean relative lesion size were determined by one-sample t-test. Levels of significance are indicated above the bars by underlined star symbols (i.e., * p < 0.05, ** p < 0.01).

4. Discussion Current practices to limit fungal diseases on plants rely heavily on synthetic chemical

compounds but their constant use poses a threat to the environment and to human health, and further increases the risk of pathogen resistance development [44]. Host resistance can provide effective control of pathogens but may be limited in some plant species, and pathogens can often overcome resistance [45]. Therefore, alternative methods for disease control are urgently needed to meet the increasing demands in crop yields and quality of produce [44,46,47].

In this study, we exploited the systemic properties of dsRNAs and successfully uti-lized bacteria as an alternative to HIGS and SIGS methods to deliver RNAi to filamentous fungi. Bacteria have the advantage that they can be cultured in large volumes relatively inexpensively and protect the produced dsRNAs within their cells until their application [48,49]. Our results showed that even after bacterial lysis, the dsRNAs remained resistant

Figure 5. The HT115/BcSAS1Ind strain induces RNAi in B. cinerea and a reduction in fungal growth under in vivo conditions.(A) B. cinerea inoculations on N. benthamiana leaves. Spores of the fungus were spot inoculated on leaves of N. benthamianaand 6 h later, 10 µL of the living bacteria or the whole-cell autolysates, obtained from the OD600 1.0 culture, were applied tothe same inoculation spots. All measurements were taken 72 h later. (B,C) BcSAS1 expression levels (Panel B) and the relativeinfection lesion size produced by B. cinerea in vivo on leaves of N. benthamiana (Panel C), following its exposure to living(Treatment A) or lysed (Treatment B) E. coli HT115(DE3) bacteria producing dsRNAs against BcSAS1 (HT115/BcSAS1Ind;test application) or eGFP (HT115/eGFPInd; control application). In Panels B and C, BcSAS1 expression levels and the meanrelative lesion size of the infection areas in the test applications (blue bars and yellow bars, respectively) are relative tothe control ones (grey bars, set to 1.0 and 100%, respectively). Error bars indicate standard deviations (SD) obtained fromat least three biological replicates. Significant differences in gene expression between control and test applications weredetermined by a one-way ANOVA followed by Tukey’s HSD test, whereas significant differences in the mean relative lesionsize were determined by one-sample t-test. Levels of significance are indicated above the bars by underlined star symbols(i.e., * p < 0.05, ** p < 0.01).

4. Discussion

Current practices to limit fungal diseases on plants rely heavily on synthetic chemicalcompounds but their constant use poses a threat to the environment and to human health,and further increases the risk of pathogen resistance development [44]. Host resistancecan provide effective control of pathogens but may be limited in some plant species, andpathogens can often overcome resistance [45]. Therefore, alternative methods for diseasecontrol are urgently needed to meet the increasing demands in crop yields and quality ofproduce [44,46,47].

In this study, we exploited the systemic properties of dsRNAs and successfully utilizedbacteria as an alternative to HIGS and SIGS methods to deliver RNAi to filamentous fungi.Bacteria have the advantage that they can be cultured in large volumes relatively inexpen-sively and protect the produced dsRNAs within their cells until their application [48,49].Our results showed that even after bacterial lysis, the dsRNAs remained resistant to degra-dation for at least up to 24 h, and partially until 72 h, providing enough time for thedsRNAs to be absorbed by the fungal cells and induce silencing in target genes. The use

J. Fungi 2021, 7, 125 17 of 22

of bacteria as vectors for delivering RNAi also circumvents the hurdle and limitations ofproducing and deploying transgenic plants in the field, whereas bacteria can be furtherengineered to be non-pathogenic microorganisms or to autolyze under selected inducedconditions, thus increasing the safety of application [27]. Indeed, the utility of bacteria asdelivery vehicles of RNAi has already been demonstrated against insects, nematodes, andeven mammalian cell cultures [27,29,31] but, surprisingly, it had not been tested againstfungi. We have now bridged this gap and show that bacteria that have been geneticallyengineered to produce dsRNAs against fungal genes can be used both in living form and aswhole-cell autolysates to induce RNAi in fungi. Indeed, our data showed that even a singleapplication of the recombinant bacteria was sufficient to achieve a significant reduction inproduction of aflatoxins by A. flavus or growth of B. cinerea, both under in vitro and in vivoconditions. This finding generates new opportunities for developing RNAi-based controlmethods against fungi of agronomic or medical importance by using non-pathogenic orattenuated bacteria as a dsRNAs delivery system.

In our studies, we have exploited the widely used HT115(DE3) E. coli strain as thehost bacterium for the production of dsRNAs [29], but other bacteria species could bepotentially engineered to deliver RNAi to fungi, including for example bacteria usedin biological control, plant endophytic or symbiotic bacteria, and several others [48].Indeed, symbiotic gut bacteria of insects were recently successfully engineered to de-liver RNAi to their insect hosts [50], indicating that the approach is amendable to differentbacterial species that can be used to meet the needs of specific applications. To furtherincrease the versatility of the system and make it suitable for diverse applications, weutilized two recombinant expression vectors that enabled the production of dsRNAs inHT115(DE3) under constitutive or IPTG-induced conditions. Specifically, the first vectoremployed the constitutive Kan promoter and allowed the uni-directional transcription ofthe two complementary RNA strands from the target DNA template, which then annealpost-transcriptionally in the cytoplasm to form dsRNAs. The second vector was based onthe bi-directional transcription of the target DNA sequence in a single reaction using the T7promoter, which is activated upon binding of the T7 RNA polymerase to it. T7 RNA poly-merase translation is in-terms dependent on initial IPTG-induction of transcription of theT7 gene in HT115(DE3) that encodes it. Inducible systems would generally provide higherefficiency, flexibility, and safety in use, as they enable the production and release of dsRNAsfrom bacteria under user-selected conditions. Although here we used the IPTG-activatedT7 RNA polymerase-based expression system, a range of inducible promoters that can betriggered under specific conditions could be employed [51,52]. Constitutive promoters,on the other hand, increase the ease of dsRNA production and could be used when thepersistent presence of recombinant bacteria producing dsRNAs is required to achieve adesirable and sustained silencing effect. Although the kinetics of dsRNAs productionby the two expression systems was not part of our study, both expression systems wereshown to yield sufficient amounts of biologically active dsRNAs that induced silencingin the target fungal genes. However, direct comparisons between the two systems couldnot be made, as different target sequences were cloned in the recombinant expressionvectors, whereas dsRNA production in the IPTG-activated system would vary dependingon the time of IPTG addition to the medium and its concentration [53]. Finally, althoughfrequently utilized in RNAi studies, the production of hairpin-structured RNA (hpRNA)was not included in these studies because it was demonstrated that HT115(DE3) is able toprocess hpRNA into short fragments, in spite of lacking RNaseIII activity [54].

In order to combine the benefits or trans-kingdom transfer and environmental uptakeof dsRNAs, we further engineered the HT115(DE3) E. coli strain to undergo programmedcellular autolysis induced by endolysin R, following a freeze-thaw cycle [35]. Both livingbacteria and their autolysates induced significant silencing in the target fungal genes ina bacteria concentration-dependent manner, suggesting that further increasing bacterialdensities could lead to an even higher effect. However, caution should be taken as increas-ing siRNAs concentrations past an optimum point does not necessarily lead to increased

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silencing levels and may even have an opposite effect due to saturation of the RNAi machin-ery [55,56]. Moreover, the silencing levels achieved by the two delivery methods were onpar with those obtained with external applications of purified dsRNAs, indicating that evensimple applications with living bacteria or their autolysates that bypass the purificationof dsRNAs and their biological stabilization by binding to various materials [3,28], canbe effective. However, whether one application was more effective than the other (i.e.,living bacteria or autolysates) could not be established, since a combination of parameters,including the fungal species and bacterial concentration, had an effect on the observedsilencing levels. For instance, while applications under in vitro conditions with livingbacteria were seemingly more effective in inducing AflC silencing in A. flavus, applica-tions with the bacterial autolysates were slightly more effective under in vivo conditions.When examining silencing of BcSAS1 in B. cinerea, then almost the opposite trend wasobserved, as applications under in vitro conditions of whole-cell autolysates were moreeffective at some bacterial concentrations, whereas living bacteria seem to have had a biggereffect under in vivo conditions. Such variation in the effectiveness of the two applicationmethods depending on the fungal species could perhaps relate to the way and rate bywhich the bacterially-produced dsRNAs are taken-up by the fungal cells. Although it isstill unclear how exogenous dsRNAs are internalized within fungal cells, several possiblemechanisms may be in effect, including passive diffusion through the fungal cell wall,internalization through plasma membrane-associated receptors or protein channels andpores [57], or through the endocytosis or fusion of bacterial extracellular vesicles carryingthe dsRNA [58]. It is plausible that the uptake of dsRNAs from the bacterial lysates ismediated by passive or active diffusion through the cell wall and membranes, whereasliving bacteria translocate dsRNAs to the fungal cells through the secretion of extracel-lular vesicles in a way similar to plants [59]. Indeed, several studies have demonstratedthat Gram-negative bacteria, including E. coli, release extracellular vesicles that containdiverse RNA molecules, including small RNAs (sRNAs) that can modulate gene expres-sion in recipient eukaryotic host cells [24,58,60–62]. Irrespectively of the mechanism(s)that may be in effect, differences in their efficacy and utilization among different fungalspecies could perhaps explain the lack of correlation seen between applications of living orlysed bacteria, and silencing levels achieved in A. flavus and B. cinerea under in vitro andin vivo conditions.

Finally, an increase in contact time between the fungi and the living bacteria or theirautolysates did not produce a clear benefit in terms of enhancing or maintaining silencinglevels in AflC or BcSAS1. However, one might expect such an effect, at least in applicationsof living bacteria that continuously produce dsRNAs compared with the one-time doseof dsRNAs received by the fungal cells from the bacterial autolysates. A combination ofvarious factors inherent to both the targeted genes and the fungal species could account forthis lack of a stable silencing effect over time. Among others, these may include dsRNAsabsorption rates, transcript abundance and mRNA turnover rate of the target genes, andthe presence of regulatory feedback mechanisms that could restore gene expression uponsilencing [63,64]. Indeed, upregulation of gene expression following RNAi is often reportedas a cellular response, mainly as a result of saturation of the RNAi machinery [65], whichin our case could be caused by the constant supply of dsRNAs. Nonetheless, althoughthe increase in contact time between fungi and the bacteria did not lead to a significantfurther increase in AflC or BcSAS1 silencing levels, in contrast, it had a positive effecton reducing production of aflatoxins in A. flavus and mycelial growth of B. cinerea overtime. This could possibly be due to the time needed for the inhibition of gene expressionto materialize into an observable phenotypic effect, or because silencing reduces proteinproduction at higher levels than mRNA levels, thus having a higher effect on phenotype.Indeed, discordances between mRNA downregulation and reduction in protein productionhave been documented before, thus leading to analogous disparities between changes ingene expression and phenotype severity [66,67].

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Overall, in this study, we demonstrate that genetically engineered bacteria that pro-duce dsRNAs against selected genes in fungi can be effectively used as vectors for deliver-ing RNAi to fungi with high efficiency, thus opening new avenues for disease control.

5. Conclusions

1. The E. coli HT115(DE3) strain can produce biologically active dsRNAs against targetgenes in fungi.

2. The biologically active bacterial dsRNAs were successfully delivered to fungi throughtwo different approaches: (i) treatment with living bacteria and (ii) treatment withbacterial lysates following a programmable cell autolysis step.

3. In vitro applications of dsRNA-producing bacteria show that the degree of the RNAieffect on fungi is positively correlated with the concentration of the bacteria, while thephenotypic changes (i.e., reduction in production of aflatoxins in A. flavus and reduc-tion of mycelial growth in B. cinerea) are positively correlated with the contact time.

4. In vivo applications of dsRNA-producing bacteria result in a reduction of aflatoxinsproduction by A. flavus on corn seeds and a reduction of disease symptoms caused byB. cinerea on N. benthamiana leaves.

Supplementary Materials: The following are available online at https://www.mdpi.com/2309-608X/7/2/125/s1, Figure S1: Circular maps of genetically engineered plasmids used for the productionof dsRNA in E. coli HT115(DE3). Figure S2: Bacterial growth curves of the E. coli HT115(DE3) strainstransformed with the plasmid vectors used to generate dsRNAs against AflC (HT115/AflCCon),BcSAS1 (HT115/BcSAS1Ind), or eGFP (HT115/eGFPCon and HT115/eGFPInd). Figure S3: Effect ofbacterial concentration on RNAi-induced silencing. Figure S4: Effect of contact time between fungiand bacteria on RNAi-induced silencing. Table S1: Primers used in this study. Table S2: Expressionlevels of AflC from A. flavus and BcSAS1 from B. cinerea in in vitro cultures of the fungi with livingor lysed E. coli HT115(DE3) bacteria producing dsRNAs against AflC (HT115/AflCCon) or BcSAS1(HT115/BcSAS1Ind), respectively. Table S3: Differences in expression levels of AflC from A. flavusamong in vitro cultures of the fungus with living or lysed E. coli HT115(DE3) bacteria producingdsRNAs against eGFP. Table S4: Differences in expression levels of BcSAS1 from B. cinerea amongin vitro cultures of the fungus with living or lysed E. coli HT115(DE3) bacteria producing dsRNAsagainst eGFP. Table S5: Expression levels of AflC from A. flavus and BcSAS1 from B. cinerea over time inin vitro cultures of the two fungi with living or lysed E. coli HT115(DE3) bacteria producing dsRNAsagainst AflC (HT115/AflCCon) or BcSAS1 (HT115/BcSAS1Ind), respectively. Table S6: Differencesin the expression levels of AflC from A. flavus over time among in vitro cultures of the fungus withliving or lysed E. coli HT115(DE3) bacteria producing dsRNAs against eGFP. Table S7: Amount ofaflatoxins produced by A. flavus over time in in vitro cultures of the fungus with living or lysedE. coli HT115(DE3) bacteria producing dsRNAs against AflC or eGFP. Table S8: Differences in theexpression levels of BcSAS1 from B. cinerea over time among in vitro cultures of the fungus withliving or lysed E. coli HT115(DE3) bacteria producing dsRNAs against eGFP. Table S9: Amount offungal biomass generated by B. cinerea over time in in vitro cultures of the fungus with living orlysed E. coli HT115(DE3) bacteria producing dsRNAs against BcSAS1 or eGFP. Table S10: Expressionlevels of AflC from A. flavus and the amount of aflatoxins produced by the fungus on maize kernels,following its exposure to living or lysed E. coli HT115(DE3) bacteria producing dsRNAs against AflCor eGFP. Table S11: Expression levels of BcSAS1 from B. cinerea and size of infection areas producedby the fungus on leaves of N. benthamiana, following its exposure to living or lysed E. coli HT115(DE3)bacteria producing dsRNAs against BcSAS1 or eGFP.

Author Contributions: Conceptualization, J.N.-S., L.-H.C., S.M. and I.S.; methodology, J.N.-S. andL.-H.C.; formal analysis, J.N.-S., L.-H.C., J.T.D.S., S.M. and I.S.; investigation, J.N.-S., L.-H.C., J.T.D.S.and S.M.; data curation, J.N.-S., L.-H.C. and I.S.; writing—original draft preparation, J.N.-S. and I.S.;writing—review and editing, J.N.-S. and I.S.; supervision, I.S.; project administration, I.S.; fundingacquisition, I.S. All authors have read and agreed to the published version of the manuscript.

Funding: This research was partially funded by a gift from MARS Incorporated.

Institutional Review Board Statement: Not applicable.

J. Fungi 2021, 7, 125 20 of 22

Informed Consent Statement: Not applicable.

Data Availability Statement: The data that support this study are available from the correspondingauthor upon reasonable request.

Acknowledgments: We kindly acknowledge Ali Ahmed and Richard Michelmore for fruitful sugges-tions and discussions. Anthony Salvucci is acknowledged for providing assistance during the initialphases of this research. J.T.D.S. acknowledges the funding received from the CAPES-PrInt program.

Conflicts of Interest: The authors declare no conflict of interest.

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